Abstract
In atopic dermatitis (AD), lesional skin is frequently colonized by Staphylococcus aureus, which promotes clinical symptoms of the disease. The inflammatory milieu in the skin is characterized by a Th2 response, including M2 macrophages, which cannot eradicate S. aureus. Therefore, repolarization of macrophages toward the M1 phenotype may foster control of S. aureus. Our data show that the deubiquitinating enzyme cylindromatosis (CYLD) is strongly expressed in macrophages of AD patients and prevents the clearance of S. aureus. Mechanistically, CYLD impaired M1 macrophage polarization by K63-specific deubiquitination of STAT1 and activation of the NF-κB pathway via its interaction with TRAF6, NEMO, and RIPK2. Inhibition of STAT1 and NF-κB, independently, abolished the differences between S. aureus-infected CYLD-deficient and CYLD-competent M1 macrophages. Infection of Cyld-deficient and wild-type mice with S. aureus confirmed the protective CYLD function. Collectively, our study shows that CYLD impairs the control of S. aureus in macrophages of AD patients, identifying CYLD as a potential therapeutic target.
Introduction
Eczematous skin lesions of patients with atopic dermatitis (AD) are frequently colonized with Staphylococcus aureus, and the extent of colonization increases in more severe lesions with hyperinflammation (1–5). These staphylococci contribute to disease symptoms, in particular itching and pain, by the secretion of toxins and proteases, which directly stimulate sensory nerves in the skin (6–8). In addition, these staphylococci may cause local and, in some cases, systemic infections in AD patients. The importance of the role of S. aureus in AD is supported by the fact that treatment of skin infection with antiseptics or antibiotics can improve the symptoms of AD (9).
Upon infection, control of S. aureus critically depends on protective innate immune responses. Neutrophils can phagocytose and kill the bacteria and also immobilize S. aureus through NETosis, i.e., capturing the bacteria in a dense network of DNA released by the neutrophils (10, 11). In addition, both tissue-resident and monocyte-derived macrophages contribute to the control and elimination of S. aureus (12). Macrophages can rapidly phagocytose S. aureus and activate the NF-κB pathway, which contributes to the production of anti-bacterial reactive oxygen species (ROS) and nitric oxide (13). Activation of macrophages by interferon (IFN)-γ synergizes with TLR2-mediated NF-κB activation in NO production and further enhances control of S. aureus in macrophages. These “M1-like” macrophages also contribute to the control of local S. aureus skin infection and contribute to the prevention of systemic bacterial dissemination (14). In contrast to M1 macrophages, M2-polarized macrophages have impaired anti-staphylococcal activity and capacity to control S. aureus infections (12). S. aureus can subvert these protective intracellular mechanisms to replicate and persist intracellularly in macrophages and other cell types including keratinocytes (13, 15–18).
The lesional skin of AD patients is characterized by a type 2 inflammatory milieu composed of macrophages, monocytes, B cells, CD4+, and CD8+ T cells (19, 20). This type 2 inflammatory environment may facilitate the persistence of extracellular S. aureus and also intracellularly in keratinocytes (16) and macrophages (17, 18). In AD patients, the dysregulation of the immune system also extends beyond the skin and includes altered systemic innate and adaptive immune profiles, leading to comorbidities including infections (21–23).
Immune responses are critically regulated by post-translational modifications including ubiquitination. Ubiquitination is a highly dynamic process exerted by a cascade of ubiquitin-activating E1, ubiquitin-conjugating E2, and ubiquitin E3 ligases, which can attach the ubiquitin protein to a lysine residue of a substrate protein, and by deubiquitinating enzymes (DUBs), which can cleave ubiquitin from the substrates. Ubiquitin can be attached as monomers or as ubiquitin chains to the substrates. In polyubiquitination, ubiquitin molecules are linked through any one of the internal seven lysines (K6, K11, K27, K29, K33, K48, and K63) or the N-terminal methionine residue (M1). The type of ubiquitin linkage decides on the function and fate of the substrate and can lead to proteasomal degradation by K48- and branched K11/K48-linked ubiquitin chains (24, 25) or to K63-linked ubiquitin modification of protein function including regulation of signal transduction (26).
Ubiquitination is reversible and counteracted by DUBs. The DUB cylindromatosis (CYLD) can cleave K63- and M1-linked polyubiquitin chains from substrates and regulates a broad range of key cellular processes including inflammatory responses, cell death pathways, autophagy, DNA damage, and cell proliferation (27, 28). In immune signaling, CYLD negatively regulates NF-κB by deubiquitinating TRAF2, TRAF6, TAK1, and NEMO (29–32). With respect to bacterial infections, this suppressive effect of CYLD on NF-κB prevented immunopathology in Haemophilus influenzae and Streptococcus pneumoniae infections (33, 34) but protective immune responses in Escherichia coli pneumonia and systemic listeriosis (35, 36). In addition, CYLD deubiquitinates STAT3 and RIPK2, thereby impairing protective IL-6 and NOD2-dependent protective immunity in listeriosis (37, 38).
Inactivating mutations of CYLD underlie the CYLD cutaneous syndrome, a disease characterized by the development of benign skin tumors of the hair follicles (39) including multiple familial trichoepithelioma, the Brooke–Spiegler syndrome, and familial cylindromatosis. In addition, somatic CYLD mutations have been linked to the suppression of sporadic cancers, non-alcoholic steatohepatitis (40), arterial hypertension (41), and gain-of-function mutations to neurodegenerative disorders (42) (reviewed in Marin-Rubio et al., 2023 (28)).
Although CYLD is strongly expressed in healthy skin and macrophages (28, 43), its expression and function in AD and S. aureus infections are unknown. To address these open questions, we analyzed the function of CYLD in AD patients. Our data show that CYLD is strongly expressed in the dermal macrophages and monocyte-derived macrophages (MDMs) of AD patients and that CYLD impairs STAT1- and NF-κB-dependent control of S. aureus in macrophages.
Materials and methods
Ethics statement
All animal experiments were in compliance with the German animal protection law in a protocol approved by the Landesverwaltungsamt Sachsen-Anhalt (file number: 203.h-42502-2-901, University of Magdeburg). The ethics committee of Hannover Medical School (MHH) approved the parts of the study involving patients (No. 10499-BO-K-2022).
Animals
Age- and sex-matched animals were used for the experiments. C57BL/6 wild-type (WT) mice were obtained from Janvier (Le Genest Saint Isle, France), and C57BL/6 Cyld−/− mice were kindly provided by Dr. Ramin Massoumi (Department of Laboratory Medicine, Malmö, Sweden) (44). All animals were kept under specific pathogen-free (SPF) conditions in an isolation facility at the Otto-von-Guericke University Magdeburg (Magdeburg, Germany).
Patients
Patients with AD were recruited at the Department of Dermatology and Allergy of Hannover Medical School (MHH). The work described was carried out in accordance with the Code of Ethics of the World Medical Association (Declaration of Helsinki), and patients gave their written informed consent prior to the study.
Staphylococcus aureus
Wild-type (strain SH1000) and methicillin-resistant (strain MW2) S. aureus were grown in Luria broth (LB, Oxoid, Germany), and aliquots of log-phase cultures were stored at −80°C. For infection of cells, fresh log-phase cultures were prepared from frozen stock.
THP-1 cells
The THP-1 cells (clone TIB-202) were obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA). The cells were cultured in a Roswell Park Memorial Institute (RPMI) cell culture medium supplemented with 10% fetal calf serum (FCS) and 1% penicillin/streptomycin.
Generation of CYLD-deficient THP-1 cells
CYLD was knocked down by the CRISPR/Cas9 method using SG cell line 4D-nucleofector X kit (#V4XP-4024, Lonza, Basel, Switzerland). For 1 × 106 cells, 210 pmol of duplexed gRNA was mixed with 70 pmol of Cas9 Nuclease V3 (#1081059, IDT, Coralville, IA, USA) for RNP complex synthesis, and the complex was mixed with 70 pmol of EE buffer (#1075916, IDT) to form electroporation mix. Electroporation was performed using the preset program DZ100 in the Lonza nucleofection system.
Generation of macrophages from THP-1 cells
THP-1 cells were differentiated into macrophages through Phorbol 12-myristate 13-acetate (PMA) (50 ng/mL) treatment for 48 h. The macrophages were either maintained in an unpolarized state (M0) or polarized into the M1 phenotype by stimulation with IFN-γ (20 ng/mL) and lipopolysaccharide (LPS) (10 pg/mL).
Generation of monocyte-derived macrophages
Peripheral blood mononuclear cells (PBMCs) were isolated using Ficoll density gradient centrifugation followed by magnetic cell separation of CD14+ monocytes (#480094, MojoSort, BioLegend, San Diego, CA, USA). CD14+ monocytes were then cultured for 6 days in Dulbecco’s modified Eagle medium (DMEM) medium supplemented with 50 ng macrophage colony-stimulating factor (M-CSF). Cells were harvested and stimulated as per experimental requirements. MDMs at day 7 polarized into M1 phenotype by stimulation with IFN-γ (20 ng/mL) and LPS (10 pg/mL).
In vitro infection of cells with S. aureus
In vitro infection of THP-1 macrophages and MDMs was performed as previously described (38). THP-1 macrophages and MDMs were infected with S. aureus at a multiplicity of infection (MOI) of 1:1. The dose of infection was verified by plating on Luria Broth (LB) agar. One hour post-infection, the extracellular bacteria were killed by incubating the cells with 30 µg/mL gentamicin (Sigma-Aldrich, St. Louis, MO, USA) for 30 min. The cells were thereafter washed with phosphate-buffered saline (PBS) to remove the extracellular bacteria and further cultivated in a medium containing gentamicin for the indicated time points. NF-κB was inhibited in THP-1 macrophages by treating the cells with IKK inhibitor VII (1 µM; Calbiochem, Darmstadt, Germany) for 24 h before the infection.
CFUs
The bacterial load in infected THP-1 macrophages and MDMs was enumerated as previously described (38). In brief, 24 h p.i., S. aureus-infected cells were washed twice with PBS to remove the antibiotics, the cells were then lysed with 0.1% Triton X-100, and serial dilution was made and plated on LB agar. Bacterial colonies were enumerated microscopically after incubation at 37°C for 24 h and 48 h.
Protein isolation and Western blotting
Proteins were isolated from S. aureus-infected THP-1 macrophages as previously described (38). In brief, cells were lysed using the lysis buffer (38). The lysates were centrifuged to remove the cell debris and the protein for Western blotting. For the Western blotting, equal amounts of proteins were separated on sodium dodecyl sulfate (SDS)–polyacrylamide gels as described previously (38), and the proteins were then transferred onto polyvinylidene fluoride membranes. To block the non-specific binding of antibodies, the membrane was incubated either with Blotto B [1% nonfat dry milk + 1% bovine serum albumin (BSA)], 5% nonfat dry milk, or 5% BSA for 1 h. The proteins were stained for GAPDH, phospho-STAT1 Y701, phospho-STAT1 S727, STAT-1, MyD88, IRAK-4, phospho-IRAK-4, TRAF6, NOD2, phospho-RIPK2, RIPK2, p65phospho-p65, p65, phospho-p38, p38, phospho-ERK1/2, ERK1/2, phospho-JNK, JNK, IKKγ/NEMO, CYLD, and K63-linkage-specific polyubiquitin overnight (all antibodies were obtained from Cell Signaling Technology, Frankfurt, Germany). The following day, membranes were washed using Tris-buffered saline with 0.1% Tween 20 (TBST) incubated with anti-mouse or anti-rabbit secondary antibodies (Dako, Hamburg, Germany) for 1 h. The blots were washed in TBST and developed using an ECL Plus kit (GE Healthcare, Freiburg, Germany). Western blotting (WB) images were captured using the Intas Chemo Cam Luminescent Image Analysis system® (INTAS Science Imaging Instruments, Göttingen, Germany) and analyzed using the LabImage 1D software® (Kapelan Bio-Imaging Solutions, Leipzig, Germany) (38).
Immunoprecipitation
Immunoprecipitation of CYLD and STAT1 was performed as described previously (38). In brief, proteins from uninfected and S. aureus-infected THP-1 macrophages were lysed using the lysis buffer as described. The protein samples were precleared using Gamma Bind™ G Sepharose™ beads (GE Healthcare Europe GmbH, Freiburg, Germany) to remove the proteins that non-specifically bind to the beads. The proteins were thereafter incubated with anti-STAT1 (1:100) or anti-CYLD (1:100) antibodies at 4°C overnight. IgG antibody was used as a negative control. The protein antibody complex was precipitated using fresh Gamma Bind™ G Sepharose™ beads at 4°C overnight. The protein antibody complex was then washed and incubated with 1× lane marker reducing sample buffer and heated at 99°C for 5 min. Thereafter, samples were centrifuged, and the supernatant was used to detect STAT1, CYLD, K63, TRAF6, RIPK2, IKK/NEMO, and K63-linked ubiquitin by WB. GAPDH was used as the input control (38).
Measurement of NO
The production of NO2 by S. aureus-infected THP-1 macrophages was determined using the Griess Assay Kit (Promega, Mannheim, Germany) as previously described (38). In brief, the supernatant from infected and non-infected cells was incubated first with sulfanilamide solution and thereafter with N-(1-naphthyl)ethylenediamine solution in the dark for 10 min. The concentration of NO2 was determined using a Synergy® microplate reader (Biotek, Berlin, Germany).
ROS detection
ROS in S. aureus-infected THP-1 macrophages were determined using a ROS detection kit (Enzo Life Sciences, Lörrach, Germany) as described previously (38). In brief, S. aureus-infected THP-1 macrophages were washed twice with the washing buffer. Thereafter, the cell pellet was resuspended in the detection reagent and incubated in the dark at 37°C for 30 min. The samples were analyzed by Cytek Aurora flow cytometry (Fremont, CA, USA).
LDH assay
Lactate dehydrogenase (LDH) enzyme activity was determined in S. aureus-infected THP-1 macrophages using CyQUANT LDH Cytotoxicity Assay (#C20300, Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s protocol.
CRISPR/Cas9 knockout of CYLD
Stable knockout of CYLD was generated by the CRISPR/Cas9 system using the gRNA Hs.Cas9.CYLD.1.AA:TCACTGACGGGGTGTACCAA and SG cell line 4D-nucleofector X kit (#V4XP-3024, Lonza) according to the manufacturer’s protocol.
In vitro siRNA treatment
For siRNA-mediated knockdown of STAT1, WT and CYLD−/− THP-1 macrophages were transfected with 20 nm of STAT1-specific siRNA (Dharmacon, Lafayette, CO, USA) according to the manufacturer’s instruction. Scrambled siRNA at 20 µM was used as a control. A total of 50 µL of the siRNA mixture was added to each well in a 24-well plate containing 4 × 105 macrophages and incubated at 37°C for 48 h. The efficiency of siRNA-mediated STAT1 silencing was controlled by WB.
Immunostaining
OCT-embedded tissues were cut into 6-mm cryosections, air-dried, and fixed at 4°C in acetone for 10 min. After washing, horseradish peroxidase (HRP) and Alkaline Phosphatase (AP) blocking solution was applied for 20 min (Invitrogen, Waltham, MA, USA), followed by Superblock (Thermo Scientific, Waltham, MA, USA). Rabbit polyclonal anti-CYLD (ab33929, Abcam, Cambridge, UK) or rabbit Ig (Dako X 0936, Agilent, Santa Clara, CA, USA) were applied in equal concentrations. Mouse monoclonal anti-CD3 (Agilent) was used to stain the cellular skin infiltrate. After applying the Envision+ HRP conjugate (Agilent) with anti-rabbit K4009 and anti-mouse K005, the AEC substrate kit was applied (AEC, 3-amino-9-ethylcarbazole; Zytomed Systems, Bargteheide, Germany). Images were taken using a Pannoramic MIDI II Sola (Sysmex, Norderstedt, Germany).
Single-cell RNA sequencing
Data from a previous study that included single-cell RNA sequencing were used for the investigation of CYLD expression on a single-cell level (Zhang et al., 2023). For a comprehensive explanation, refer to the corresponding methods section of the study. Briefly, skin punch biopsies were processed using the Miltenyi Biotech skin dissociation kit for subsequent cell isolation and CD2 enrichment, followed by pooling and loading onto the chromium chip (10x Genomics, Pleasanton, CA, USA). The CellRanger pipeline version 3.1.0 was employed to align reads to the human reference genome GRCH38. The expression matrix generated was then analyzed using the Seurat package version 4.2.3. UMAP and feature plots were generated using their respective functions.
Statistics
The statistical significance was determined using the software Prism 9 with respective tests as mentioned in the figure legends, and p-values of ≤0.05 were considered significant. All experiments were performed at least twice.
Results
CYLD expression in eczematous skin lesions of AD patients
The expression of CYLD in different organs and cells under homeostatic conditions has been summarized in “The Human Protein Atlas” (https://www.proteinatlas.org/ENSG00000083799-CYLD). These data show that CYLD is expressed in all organs including the skin, with strong protein and mRNA expression in keratinocytes, Langerhans cells, and immune cells including T cells, macrophages, and B cells but not in melanocytes (43). CYLD is expressed in all leukocyte populations including granulocytes and monocytes (https://www.proteinatlas.org/ENSG00000083799-CYLD/immune+cell). Transcriptome analysis of skin biopsies from 10 AD patients identified CYLD mRNA in T cells, natural killer cells (Figure 1A), and different macrophage subtypes including M2 and dendritic cells (DCs) (Figure 1B). mRNA expression in keratinocytes, fibroblasts, vascular endothelial cells, and pericytes appeared weaker but was present as well, while CYLD expression was also detectable in PBMC samples of the same donors including different myeloid cells (Figures 1C, D). Our histological analysis of CYLD expression in AD patients revealed the expression of CYLD in CD3+ T cells as well as non-T cells (Figure 1E). CYLD expression appeared particularly strong in the lesional skin of a patient with clinical signs of superinfected eczema (Figure 1E, patient AD3). Our further histological analysis confirmed the widespread CYLD expression in the cellular infiltrate and keratinocytes of inflamed AD skin lesions as compared to healthy skin (Supplementary Figure S1). Thus, CYLD is expressed in the skin of both healthy persons and AD patients with expression in identical cell types including skin macrophages and monocytes in the blood.
Figure 1
Increased CYLD expression and impaired control of S. aureus in monocyte-derived macrophages of AD patients
Since AD is characterized by a type 2 inflammation including M2 macrophage polarization (19) and activation of the IFN-γ/STAT1 and TLR2/4 pathways synergize in the control of S. aureus, we first determined whether M1 polarization of macrophages improves the control of S. aureus-infected macrophages of AD patients to the same extent as in healthy controls. In these experiments, we isolated CD14+ monocytes from the blood of AD patients and healthy controls, and we differentiated these cells into macrophages by M-CSF treatment followed by IFN-γ/LPS treatment. Control of S. aureus was significantly impaired in MDMs from all investigated AD patients (Figure 2A). Since CYLD can impair the cell-intrinsic control of intracellular bacteria (37, 38), we determined the CYLD expression in the infected MDMs. WB analysis showed that the impaired capacity of M1-polarized MDMs of AD correlated with an increased CYLD protein expression in these infected cells (Figure 2B).
Figure 2
Thus, AD patients are characterized by an overexpression of CYLD in macrophages and impaired control of S. aureus. To corroborate the inhibitory function of CYLD on the control of S. aureus in M1-polarized macrophages of AD patients, we deleted CYLD by siRNA in IFN-γ/LPS-primed (M1) macrophages of healthy blood donors and AD patients (Figure 2C). Upon infection, the control of S. aureus was significantly improved in CYLD-deleted macrophages of AD patients as compared to control siRNA-treated macrophages (Figure 2D). Thus, CYLD is upregulated in MDMs of AD patients and impairs the control of S. aureus in M1-polarized MDMs, demonstrating that CYLD is a macrophage intrinsic inhibitor of the control of S. aureus.
CYLD inhibits M1 macrophage polarization, production of anti-bacterial reactive oxygen species, and control of S. aureus
To determine the mechanisms of how CYLD impairs intrinsic immunity to S. aureus, we established CYLD-deficient THP-1 monocytes by CRISPR/Cas9 (Figure 3A). Upon PMA-mediated differentiation into macrophages and subsequent IFN-γ/LPS stimulation, CYLD-deficient THP-1 cells expressed higher levels of CD80, a marker for M1 macrophages, and TNF and IL-6 mRNA, two prototypic M1 macrophage cytokines important for the control of S. aureus (Figure 3B). In IFN-γ/LPS-stimulated THP-1 macrophages, CYLD deletion significantly improved control of methicillin-sensitive and methicillin-resistant S. aureus, whereas no differences between the two genotypes were observed for unstimulated macrophages (Figures 3C, D). Analysis of anti-bacterial nitric oxide and reactive oxygen species revealed that ROS were significantly increased in infected CYLD-deficient THP-1 24 h p.i., whereas NO levels did not differ (Figures 3E–G). A kinetic analysis of bacterial loads revealed that CYLD does not influence the entry of S. aureus into the macrophages (Figure 3H) but impairs intracellular bacterial control, which leads to the death of macrophages (Figure 3I). These data illustrate that CYLD impaired M1 macrophage differentiation, control of S. aureus, and ROS production.
Figure 3
CYLD inhibits STAT1, MyD88, and NOD2 signaling in S. aureus-infected M1-polarized macrophages
Since M1-polarized macrophages showed improved control of S. aureus and anti-bacterial activity in CYLD-deficient macrophages, we determined the impact of CYLD on i) IFN-γ-induced STAT1 activation, ii) the effect of LPS (TLR4) and S. aureus (TLR2)-induced MyD88 signaling, and iii) S. aureus-activated NOD2 pathways.
WB analyses showed that CYLD impaired STAT1 phosphorylation at tyrosine 701 and serine 727, which cooperatively led to the induction of IFN-γ-induced genes (Sadzak et al., 2008; Varinou et al., 2003) (Figure 4A). Co-immunoprecipitation experiments newly identified that CYLD directly interacted with STAT1 (Figures 4B, C). STAT1 immunoprecipitation and subsequent WB analysis of K63 ubiquitin revealed that S. aureus infection induced increased K63 ubiquitination of STAT1 in CYLD-deficient macrophages as compared to WT macrophages at 2 h p.i. (Figure 4D). In agreement with published data in other models (29, 30, 32, 38), CYLD also interacted with TRAF6, IKKγ/NEMO, and RIPK2 in M1-polarized S. aureus-infected THP-1 macrophages (Figure 4E). This was associated with impaired MyD88 and NOD2/RIPK2 signaling, leading to reduced downstream NF-κB activation shown by impaired phosphorylation of p65 in S. aureus-infected CYLD-competent macrophages (Figure 4F). Thus, CYLD inhibits simultaneously key pathways leading to reduced activation of the transcription factors NF-κB and STAT1.
Figure 4
The enhanced control of intracellular S. aureus in CYLD-deficient THP-1-derived M1 macrophages is NF-κB- and STAT1-dependent
To determine the functional importance of CYLD-regulated NF-κB and STAT1 activation for the control of S. aureus, we stimulated THP-1 macrophages with IFN-γ/LPS for M1 polarization and treated the cells prior to infection with the IKK VII inhibitor to inhibit activation of NF-κB and with STAT1 siRNA to inhibit STAT1. Inhibition of NF-κB activation and STAT1 resulted in a strong increase of colony-forming units (CFUs) of S. aureus in both WT and CYLD-deficient THP-1 macrophages, and both treatments abolished the differences between the two genotypes (Figures 5A, B). Thus, the inhibition of both STAT1 and NF-κB by CYLD is critical for the control of S. aureus in macrophages.
Figure 5
Cyld-deficient mice are protected from S. aureus infection
The data presented identify that CYLD is increasingly expressed in macrophages of AD patients and inhibits the control of S. aureus in human macrophages. To further validate an inhibitory function of CYLD in S. aureus infection and to evaluate whether systemic CYLD inhibition may be a therapeutic option to ameliorate S. aureus infection, we infected Cyld-deficient and WT mice with S. aureus. Cyld-deficient mice had significantly reduced weight loss (Figure 6A) and improved pathogen control in the liver and kidney at the acute and chronic stages of infection (Figures 6B, C). In good agreement with the human macrophages, CYLD-deficient murine bone marrow-derived macrophages showed better clearance of S. aureus compared to WT macrophages (Figure 6D). This qualifies CYLD as a relevant therapeutic target to ameliorate S. aureus infections.
Figure 6
Discussion
Macrophages play a fundamental role in the control of local and systemic infections with S. aureus (12, 13, 45). In murine S. aureus skin infection, M1-polarized macrophages contribute to the control of the infection, whereas M2-polarized macrophages are associated with impaired killing of S. aureus and spread of the pathogen (46). In AD, the cutaneous inflammatory lesions are characterized by a type 2 inflammatory milieu including M2-polarized macrophages infiltrating the diseased skin (20, 47, 48). A shift to the M1 phenotype of macrophages would reduce S. aureus colonization and infections in AD patients. However, this study illustrates that M1-polarized MDMs of AD patients express an increased amount of CYLD and have an impaired capacity to kill S. aureus. The functional importance of CYLD is shown in THP-1 macrophages, which only incompletely shift to an M1 phenotype as illustrated by reduced CD80 expression and TNF production and by the improved control of S. aureus in CYLD-deficient M1-polarized MDMs of healthy blood donors and THP-1 cells.
The present study newly identifies that CYLD binds to STAT1 upon S. aureus infection of M1-polarized human MDMs. This interaction resulted in a CYLD-dependent reduction of K63 polyubiquitination of STAT1 and an impaired STAT1 phosphorylation. CYLD inhibited both the phosphorylation at STAT1 (Y701), which is important for nuclear accumulation of the STAT1 transcription factor, and the subsequent nuclear phosphorylation at STAT1 (S727), which is required for gene transcription (49, 50). The functional importance of STAT1 for the control of S. aureus was illustrated by the impaired control of S. aureus in M1-polarized MDMs upon siRNA-mediated inhibition of STAT1.
The induction of an anti-staphylococcal function of macrophages also requires the engagement of pathogen pattern recognition receptors leading to the activation of NF-κB (12). In the present study, the pre-stimulation with the TLR4 agonist LPS and the subsequent S. aureus infection activating TLR2 and NOD2 by lipoteichoic acid and muramyl dipeptide led to the activation of NF-κB and NF-κB-dependent production of anti-bacterial ROS. Production of ROS is important and has a superior role in comparison to NO in controlling S. aureus infections (13, 51–55). In agreement with previous studies, we detected an interaction of CYLD with RIPK2, TRAF6, and IKKγ/NEMO, which are critical signaling molecules in the NOD2 and TLR2/4 pathways leading to NF-κB activation (29, 30, 32, 38). CYLD inhibits the activation of these signaling molecules by its K63 deubiquitinating activity. The key role of CYLD-mediated NF-κB inhibition as a critical factor impairing control of S. aureus in macrophages is proven by the strong increase and abolishment of differences in CFUs between CYLD-competent and CYLD-deficient macrophages. Since the numbers of S. aureus were identical in both NF-κB-inhibited and STAT1 siRNA-treated WT and CYLD−/− macrophages, the two pathways determine independently from each other the control of S. aureus and cannot compensate for each other.
Of note, infection with S. aureus led to a CYLD-independent and equal activation of MAP kinases including c-Jun-N-terminal kinase (JNK), p38, and ERK in CYLD-competent and CYLD-deficient MDMs. Previously, we have shown that the inhibition of RIPK2 by CYLD impairs the activation of MAP kinases and ERK1/2-induced autophagy in Listeria monocytogenes-infected macrophages (38). In the present study, MAP kinases, in particular JNK, were equally activated in CYLD-deficient and CYLD-competent MDMs. This difference may be explained by the manipulation of MAP kinases by S. aureus, which may undermine JNK function by a yet unresolved mechanism to induce its persistence in macrophages (56). The pathogen S. aureus has been found to prominently colonize and thereby overgrow commensal microbiota on the skin of AD patients, aggravating AD on different levels. Its role in the disease is further underlined by the finding that coagulase-negative staphylococci, which are able to suppress the growth of S. aureus, were less present on the skin of babies who had developed AD at the second study visit after 1 year (57). Healthy mice treated with S. aureus strains derived from AD patients develop a robust immune reaction including type 2 cytokines and induction of skin-resident T cells (58). While keratinocytes act as sensors of S. aureus infection (59), S. aureus is capable of infiltrating keratinocytes and immune cells (58, 60), thereby reducing the effectivity of antimicrobial or antibiotic treatment. Immune cells, which are crucial for the final clearance, are efficiently repressed by evasion strategies (17). Strengthening the hosts’ cellular mechanism to control S. aureus infection could therefore be a potent therapy option in AD.
Our data show that M1 polarization of macrophages augments the anti-staphylococcal activity of human and murine macrophages. Unleashing the CYLD break on NF-κB further enhances the control of S. aureus of M1-polarized macrophages. The in vivo data illustrating that Cyld−/− mice have an improved course and control of S. aureus in acute and chronic systemic infection further corroborate that CYLD is a potential therapeutic target to reduce S. aureus colonization and infection.
The potential of CYLD inhibition in infectious diseases
CYLD is known to negatively regulate various signaling pathways, including NF-κB (61, 62), MAPK (38), and JAK/STAT (37, 38, 63) pathways, which are crucial for the inflammatory response. Our previous studies have identified CYLD as an inhibitor of immune responses during bacterial (37, 38) and parasitic infections (64), and inhibition of CYLD resulted in a positive disease outcome. Inhibition of CYLD in hepatocytes promoted IL-6/STAT3-mediated fibrin production and prevented the spread of L. monocytogenes (37). Ablation of CYLD in macrophages enhanced NF-κB-mediated production of reactive oxygen species, resulting in improved clearance of L. monocytogenes (38). Depletion of CYLD in T cells attenuates T-cell responses, prevents blood–brain barrier disruption, and protects mice from experimental cerebral malaria (64). Furthermore, Lim et al. (2007) showed that CYLD deficiency protected mice from S. pneumoniae pneumolysin (PLY)-induced Acute lung injury (ALI) and lethality (34). In the current study, we identified CYLD as an inhibitor of the anti-S. aureus immune response in human and murine macrophages. These studies highlight the potential of CYLD as a therapeutic target in infectious diseases. However, it should be noted that CYLD negatively regulates NF-κB-dependent inflammation during non-typeable H. influenzae infection and protects mice from deleterious inflammatory responses (65). Similarly, CYLD impairs inflammation and protects mice from E. coli-induced pneumonia (35), suggesting that the protective or detrimental function of CYLD in infectious diseases depends on the underlying infection.
Impact on macrophage polarization
M1 macrophages are crucial in defending against pathogens through phagocytosis and the secretion of pro-inflammatory cytokines. Several bacteria, including S. aureus (66), Mycobacterium (67), Salmonella (68), and Coxiella (69), promote M2 polarization to evade the pro-inflammatory response and enhance their survival within macrophages. Ablation of CYLD would foster M1 polarization, enhancing their antimicrobial activities and leading to improved outcomes in these bacterial infections.
Therapeutic strategies
Commercially available small molecule inhibitors that target the USP family of deubiquitinases, such as NSC-687852, NSC112200, and WP1130, could be used to inhibit CYLD. More specifically, subquinocin, a small molecule inhibitor primarily targeting CYLD, could serve as a potential therapeutic strategy against infectious diseases (70). Additionally, proteolysis-targeting chimeras (PROTACs), which utilize the ubiquitin–proteasome system to target specific proteins for degradation (71), could be used to specifically target CYLD for proteasomal degradation, making it a promising therapeutic option.
Statements
Data availability statement
The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found below: EGAD00001010106 (EGA; https://ega-archive.org).
Ethics statement
The ethics committee of Hannover Medical School (MHH) approved the parts of the study involving patients (No. 10499-BO-K-2022). The studies were conducted in accordance with the local legislation and institutional requirements. The participants provided their written informed consent to participate in this study. The animal study was approved by Landesverwaltungsamt Sachsen-Anhalt (file number: 203.h-42502-2-901, University of Magdeburg). The study was conducted in accordance with the local legislation and institutional requirements.
Author contributions
CS: Formal analysis, Investigation, Methodology, Validation, Writing – review & editing. KH: Formal analysis, Investigation, Methodology, Validation, Writing – review & editing. ST: Formal analysis, Investigation, Methodology, Validation, Writing – review & editing. MN: Formal analysis, Validation, Writing – review & editing. TW: Conceptualization, Writing – review & editing. LR: Writing – review & editing, Investigation, Methodology, Validation, Writing – original draft. GN: Validation, Writing – original draft, Writing – review & editing, Conceptualization, Supervision. DS: Conceptualization, Funding acquisition, Supervision, Writing – original draft, Writing – review & editing.
Funding
The author(s) declare financial support was received for the research, authorship, and/or publication of this article. This work was funded by the German Research Foundation, Excellence Strategy—EXC 2155 “RESIST”—Project number 390874280.
Acknowledgments
We thank Birgit Brenneke, Kerstin Ellrott, and Gabriele Begemann for expert technical assistance. Further, we thank all volunteers for their contribution to this project.
Conflict of interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Generative AI statement
The author(s) declare that no Generative AI was used in the creation of this manuscript.
Publisher’s note
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.
Supplementary material
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fimmu.2025.1507989/full#supplementary-material
Supplementary Figure 1Increased CYLD protein expression in the epidermal skin of atopic dermatitis patients. Exemplary pictures of CYLD protein expression (brown, AEC) in the skin of three patients with AD and one healthy control. Sections were immunostained with a polyclonal rabbit anti-CYLD antibody or a respective isotype control and counterstained with hemalum. Scale bar = 100µm.
References
1
EdslevSMOlesenCMNorresletLBInghamACIversenSLiljeBet al. Staphylococcal communities on skin are associated with atopic dermatitis and disease severity. Microorganisms. (2021) 9. doi: 10.3390/microorganisms9020432
2
GuzikTJBzowskaMKasprowiczACzerniawska-MysikGWojcikKSzmydDet al. Persistent skin colonization with Staphylococcus aureus in atopic dermatitis: relationship to clinical and immunological parameters. Clin Exp Allergy. (2005) 35:448–55. doi: 10.1111/j.1365-2222.2005.02210.x
3
KongHHOhJDemingCConlanSGriceEABeatsonMAet al. Temporal shifts in the skin microbiome associated with disease flares and treatment in children with atopic dermatitis. Genome Res. (2012) 22:850–9. doi: 10.1101/gr.131029.111
4
Saheb KashafSHarkinsCPDemingCJoglekarPConlanSHolmesCJet al. Staphylococcal diversity in atopic dermatitis from an individual to a global scale. Cell Host Microbe. (2023) 31:578–592 e576. doi: 10.1016/j.chom.2023.03.010
5
WichmannKUterWWeissJBreuerKHeratizadehAMaiUet al. Isolation of alpha-toxin-producing Staphylococcus aureus from the skin of highly sensitized adult patients with severe atopic dermatitis. Br J Dermatol. (2009) 161:300–5. doi: 10.1111/j.1365-2133.2009.09229.x
6
DengLCostaFBlakeKJChoiSChandrabalanAYousufMSet al. S. aureus drives itch and scratch-induced skin damage through a V8 protease-PAR1 axis. Cell. (2023) 186:5375–5393 e5325. doi: 10.1016/j.cell.2023.10.019
7
GeogheganJAIrvineADFosterTJ. Staphylococcus aureus and atopic dermatitis: A complex and evolving relationship. Trends Microbiol. (2018) 26:484–97. doi: 10.1016/j.tim.2017.11.008
8
LangerKBreuerKKappAWerfelT. Staphylococcus aureus-derived enterotoxins enhance house dust mite-induced patch test reactions in atopic dermatitis. Exp Dermatol. (2007) 16:124–9. doi: 10.1111/j.1600-0625.2006.00523.x
9
BreuerKSHAKappAWerfelT. Staphylococcus aureus: colonizing features and influence of an antibacterial treatment in adults with atopic dermatitis. Br J Dermatol. (2002) 147:55–61. doi: 10.1046/j.1365-2133.2002.04872.x
10
SpaanANSurewaardBGNijlandRvan StrijpJA. Neutrophils versus Staphylococcus aureus: a biological tug of war. Annu Rev Microbiol. (2013) 67:629–50. doi: 10.1146/annurev-micro-092412-155746
11
von Kockritz-BlickwedeMWinstelV. Molecular prerequisites for neutrophil extracellular trap formation and evasion mechanisms of staphylococcus aureus. Front Immunol. (2022) 13:836278. doi: 10.3389/fimmu.2022.836278
12
PidwillGRGibsonJFColeJRenshawSAFosterSJ. The role of macrophages in staphylococcus aureus infection. Front Immunol. (2020) 11:620339. doi: 10.3389/fimmu.2020.620339
13
SurewaardBGDenisetJFZempFJAmreinMOttoMConlyJet al. Identification and treatment of the Staphylococcus aureus reservoir in vivo. J Exp Med. (2016) 213:1141–51. doi: 10.1084/jem.20160334
14
ChanLCRossettiMMillerLSFillerSGJohnsonCWLeeHKet al. Protective immunity in recurrent Staphylococcus aureus infection reflects localized immune signatures and macrophage-conferred memory. Proc Natl Acad Sci U.S.A. (2018) 115:E11111–9. doi: 10.1073/pnas.1808353115
15
GreshamHDLowranceJHCaverTEWilsonBSCheungALLindbergFP. Survival of Staphylococcus aureus inside neutrophils contributes to infection. J Immunol. (2000) 164:3713–22. doi: 10.4049/jimmunol.164.7.3713
16
SoongGPaulinoFWachtelSParkerDWickershamMZhangDet al. Methicillin-resistant Staphylococcus aureus adaptation to human keratinocytes. mBio. (2015) 6. doi: 10.1128/mBio.00289-15
17
ThammavongsaVKimHKMissiakasDSchneewindO. Staphylococcal manipulation of host immune responses. Nat Rev Microbiol. (2015) 13:529–43. doi: 10.1038/nrmicro3521
18
TuchscherrLMedinaEHussainMVolkerWHeitmannVNiemannSet al. Staphylococcus aureus phenotype switching: an effective bacterial strategy to escape host immune response and establish a chronic infection. EMBO Mol Med. (2011) 3:129–41. doi: 10.1002/emmm.201000115
19
BeckLACorkMJAmagaiMDe BenedettoAKabashimaKHamiltonJDet al. Type 2 inflammation contributes to skin barrier dysfunction in atopic dermatitis. JID Innov. (2022) 2:100131. doi: 10.1016/j.xjidi.2022.100131
20
ZhangBRoesnerLMTraidlSKoekenVXuCJWerfelTet al. Single-cell profiles reveal distinctive immune response in atopic dermatitis in contrast to psoriasis. Allergy. (2023) 78:439–53. doi: 10.1111/all.15486
21
BrunnerPMSilverbergJIGuttman-YasskyEPallerASKabashimaKAmagaiMet al. Increasing comorbidities suggest that atopic dermatitis is a systemic disorder. J Invest Dermatol. (2017) 137:18–25. doi: 10.1016/j.jid.2016.08.022
22
DroitcourtCVittrupIKerbratSEgebergAThyssenJP. Risk of systemic infections in adults with atopic dermatitis: A nationwide cohort study. J Am Acad Dermatol. (2021) 84:290–9. doi: 10.1016/j.jaad.2020.07.111
23
JinSPLeeKBangYJJeonYHJungSChoiSJet al. Mapping the immune cell landscape of severe atopic dermatitis by single-cell RNA-seq. Allergy. (2024) 79:1584–97. doi: 10.1111/all.16121
24
ChauVTobiasJWBachmairAMarriottDEckerDJGondaDKet al. A multiubiquitin chain is confined to specific lysine in a targeted short-lived protein. Science. (1989) 243:1576–83. doi: 10.1126/science.2538923
25
MeyerHJRapeM. Enhanced protein degradation by branched ubiquitin chains. Cell. (2014) 157:910–21. doi: 10.1016/j.cell.2014.03.037
26
RahmanSWolbergerC. Breaking the K48-chain: linking ubiquitin beyond protein degradation. Nat Struct Mol Biol. (2024) 31:216–8. doi: 10.1038/s41594-024-01221-w
27
KomanderDReyes-TurcuFLicchesiJDOdenwaelderPWilkinsonKDBarfordD. Molecular discrimination of structurally equivalent Lys 63-linked and linear polyubiquitin chains. EMBO Rep. (2009) 10:466–73. doi: 10.1038/embor.2009.55
28
Marin-RubioJLRaoteIInnsJDobson-StoneCRajanN. CYLD in health and disease. Dis Model Mech. (2023) 16. doi: 10.1242/dmm.050093
29
BrummelkampTRNijmanSMDiracAMBernardsR. Loss of the cylindromatosis tumour suppressor inhibits apoptosis by activating NF-kappaB. Nature. (2003) 424:797–801. doi: 10.1038/nature01811
30
KovalenkoAChable-BessiaCCantarellaGIsraelAWallachDCourtoisG. The tumour suppressor CYLD negatively regulates NF-kappaB signalling by deubiquitination. Nature. (2003) 424:801–5. doi: 10.1038/nature01802
31
ReileyWWJinWLeeAJWrightAWuXTewaltEFet al. Deubiquitinating enzyme CYLD negatively regulates the ubiquitin-dependent kinase Tak1 and prevents abnormal T cell responses. J Exp Med. (2007) 204:1475–85. doi: 10.1084/jem.20062694
32
TrompoukiEHatzivassiliouETsichritzisTFarmerHAshworthAMosialosG. CYLD is a deubiquitinating enzyme that negatively regulates NF-kappaB activation by TNFR family members. Nature. (2003) 424:793–6. doi: 10.1038/nature01803
33
LimJHJonoHKogaTWooCHIshinagaHBournePet al. Tumor suppressor CYLD acts as a negative regulator for non-typeable Haemophilus influenza-induced inflammation in the middle ear and lung of mice. PloS One. (2007) 2:e1032. doi: 10.1371/journal.pone.0001032
34
LimJHStirlingBDerryJKogaTJonoHWooCHet al. Tumor suppressor CYLD regulates acute lung injury in lethal Streptococcus pneumoniae infections. Immunity. (2007) 27:349–60. doi: 10.1016/j.immuni.2007.07.011
35
LimJHHaUHWooCHXuHLiJD. CYLD is a crucial negative regulator of innate immune response in Escherichia coli pneumonia. Cell Microbiol. (2008) 10:2247–56. doi: 10.1111/j.1462-5822.2008.01204.x
36
WurmRJustSWangXWexKSchmidUBlanchardNet al. Protective dendritic cell responses against listeriosis induced by the short form of the deubiquitinating enzyme CYLD are inhibited by full-length CYLD. Eur J Immunol. (2015) 45:1366–76. doi: 10.1002/eji.201445116
37
NishanthGDeckertMWexKMassoumiRSchweitzerKNaumannMet al. CYLD enhances severe listeriosis by impairing IL-6/STAT3-dependent fibrin production. PloS Pathog. (2013) 9:e1003455. doi: 10.1371/journal.ppat.1003455
38
WexKSchmidUJustSWangXWurmRNaumannMet al. Receptor-interacting protein kinase-2 inhibition by CYLD impairs antibacterial immune responses in macrophages. Front Immunol. (2015) 6:650. doi: 10.3389/fimmu.2015.00650
39
NagyNDuboisASzellMRajanN. Genetic testing in CYLD cutaneous syndrome: an update. Appl Clin Genet. (2021) 14:427–44. doi: 10.2147/TACG.S288274
40
JiYXHuangZYangXWangXZhaoLPWangPXet al. The deubiquitinating enzyme cylindromatosis mitigates nonalcoholic steatohepatitis. Nat Med. (2018) 24:213–23. doi: 10.1038/nm.4461
41
ZhouJJLiHLiLLiYWangPHMengXMet al. CYLD mediates human pulmonary artery smooth muscle cell dysfunction in congenital heart disease-associated pulmonary arterial hypertension. J Cell Physiol. (2021) 236:6297–311. doi: 10.1002/jcp.30298
42
Dobson-StoneCHalluppMShahheydariHRagagninAMGChattertonZCarew-JonesFet al. CYLD is a causative gene for frontotemporal dementia - amyotrophic lateral sclerosis. Brain. (2020) 143:783–99. doi: 10.1093/brain/awaa039
43
UhlenMOksvoldPFagerbergLLundbergEJonassonKForsbergMet al. Towards a knowledge-based human protein atlas. Nat Biotechnol. (2010) 28:1248–50. doi: 10.1038/nbt1210-1248
44
MassoumiRChmielarskaKHenneckeKPfeiferAFasslerR. Cyld inhibits tumor cell proliferation by blocking Bcl-3-dependent NF-kappaB signaling. Cell. (2006) 125:665–77. doi: 10.1016/j.cell.2006.03.041
45
YajjalaVKThomasVCBauerCScherrTDFischerKJFeyPDet al. Resistance to acute macrophage killing promotes airway fitness of prevalent community-acquired staphylococcus aureus strains. J Immunol. (2016) 196:4196–203. doi: 10.4049/jimmunol.1600081
46
AsaiATsudaYKobayashiMHanafusaTHerndonDNSuzukiF. Pathogenic role of macrophages in intradermal infection of methicillin-resistant Staphylococcus aureus in thermally injured mice. Infect Immun. (2010) 78:4311–9. doi: 10.1128/IAI.00642-10
47
HashimotoTYokozekiHKarasuyamaHSatohT. IL-31-generating network in atopic dermatitis comprising macrophages, basophils, thymic stromal lymphopoietin, and periostin. J Allergy Clin Immunol. (2023) 151:737–746 e736. doi: 10.1016/j.jaci.2022.11.009
48
KasraieSWerfelT. Role of macrophages in the pathogenesis of atopic dermatitis. Mediators Inflammation. (2013), 942375. doi: 10.1155/2013/942375
49
SadzakISchiffMGattermeierIGlinitzerRSauerISaalmullerAet al. Recruitment of Stat1 to chromatin is required for interferon-induced serine phosphorylation of Stat1 transactivation domain. Proc Natl Acad Sci U.S.A. (2008) 105:8944–9. doi: 10.1073/pnas.0801794105
50
VarinouLRamsauerKKaraghiosoffMKolbeTPfefferKMullerMet al. Phosphorylation of the Stat1 transactivation domain is required for full-fledged IFN-gamma-dependent innate immunity. Immunity. (2003) 19:793–802. doi: 10.1016/s1074-7613(03)00322-4
51
HashimotoMTawaratsumidaKKariyaHAoyamaKTamuraTSudaY. Lipoprotein is a predominant Toll-like receptor 2 ligand in Staphylococcus aureus cell wall components. Int Immunol. (2006) 18:355–62. doi: 10.1093/intimm/dxh374
52
PizzollaAHultqvistMNilsonBGrimmMJEneljungTJonssonIMet al. Reactive oxygen species produced by the NADPH oxidase 2 complex in monocytes protect mice from bacterial infections. J Immunol. (2012) 188:5003–11. doi: 10.4049/jimmunol.1103430
53
SchafflerHDemirciogluDDKuhnerDMenzSBenderAAutenriethIBet al. NOD2 stimulation by Staphylococcus aureus-derived peptidoglycan is boosted by Toll-like receptor 2 costimulation with lipoproteins in dendritic cells. Infect Immun. (2014) 82:4681–8. doi: 10.1128/IAI.02043-14
54
TakeuchiOHoshinoKKawaiTSanjoHTakadaHOgawaTet al. Differential roles of TLR2 and TLR4 in recognition of gram-negative and gram-positive bacterial cell wall components. Immunity. (1999) 11:443–51. doi: 10.1016/s1074-7613(00)80119-3
55
TakeuchiOTakedaKHoshinoKAdachiOOgawaTAkiraS. Cellular responses to bacterial cell wall components are mediated through MyD88-dependent signaling cascades. Int Immunol. (2000) 12:113–7. doi: 10.1093/intimm/12.1.113
56
WatanabeIIchikiMShiratsuchiANakanishiY. TLR2-mediated survival of Staphylococcus aureus in macrophages: a novel bacterial strategy against host innate immunity. J Immunol. (2007) 178:4917–25. doi: 10.4049/jimmunol.178.8.4917
57
KennedyEAConnollyJHourihaneJOFallonPGMcLeanWHIMurrayDet al. Skin microbiome before development of atopic dermatitis: Early colonization with commensal staphylococci at 2 months is associated with a lower risk of atopic dermatitis at 1 year. J Allergy Clin Immunol. (2017) 139:166–72. doi: 10.1016/j.jaci.2016.07.029
58
BraunCBadiouCGuironnet-PaquetAIwataMLeniefVMosnierAet al. Staphylococcus aureus-specific skin resident memory T cells protect against bacteria colonization but exacerbate atopic dermatitis-like flares in mice. J Allergy Clin Immunol. (2024) 154:355–74. doi: 10.1016/j.jaci.2024.03.032
59
BitscharKWolzCKrismerBPeschelASchittekB. Keratinocytes as sensors and central players in the immune defense against Staphylococcus aureus in the skin. J Dermatol Sci. (2017) 87:215–20. doi: 10.1016/j.jdermsci.2017.06.003
60
KintarakSWhawellSASpeightPMPackerSNairSP. Internalization of Staphylococcus aureus by human keratinocytes. Infect Immun. (2004) 72:5668–75. doi: 10.1128/IAI.72.10.5668-5675.2004
61
TrompoukiEHatzivassiliouETsichritzisTFarmerHAshworthAMosialosG. CYLD is a deubiquitinating enzyme that negatively regulates NF-κB activation by TNFR family members. Nature. (2003) 424:793–6. doi: 10.1038/nature01803
62
JonoHLimJHChenLFXuHTrompoukiEPanZKet al. NF-kappaB is essential for induction of CYLD, the negative regulator of NF-kappaB: evidence for a novel inducible autoregulatory feedback pathway. J Biol Chem. (2004) 279:36171–4. doi: 10.1074/jbc.M406638200
63
HuyenNTNgocNTGiangNHTrangDTHanhHHBinhVDet al. CYLD stimulates macrophage phagocytosis of leukemic cells through STAT1 signalling in acute myeloid leukemia. PloS One. (2023) 18:e0283586. doi: 10.1371/journal.pone.0283586
64
SchmidUStenzelWKoschelJRaptakiMWangXNaumannMet al. The deubiquitinating enzyme cylindromatosis dampens CD8(+) T cell responses and is a critical factor for experimental cerebral malaria and blood-brain barrier damage. Front Immunol. (2017) 8:27. doi: 10.3389/fimmu.2017.00027
65
WangWYKomatsuKHuangYWuJZhangWLeeJYet al. CYLD negatively regulates nontypeable Haemophilus influenzae-induced IL-8 expression via phosphatase MKP-1-dependent inhibition of ERK. PloS One. (2014) 9:e112516. doi: 10.1371/journal.pone.0112516
66
XuFKangYZhangHPiaoZYinHDiaoRet al. Akt1-mediated regulation of macrophage polarization in a murine model of Staphylococcus aureus pulmonary infection. J Infect Dis. (2013) 208:528–38. doi: 10.1093/infdis/jit177
67
ShaSShiYTangYJiaLHanXLiuYet al. Mycobacterium tuberculosis Rv1987 protein induces M2 polarization of macrophages through activating the PI3K/Akt1/mTOR signaling pathway. Immunol Cell Biol. (2021) 99:570–85. doi: 10.1111/imcb.12436
68
LeibaJSipkaTBegon-PesciaCBernardelloMTairiSBossiLet al. Dynamics of macrophage polarization support Salmonella persistence in a whole living organism. Elife. (2024) 13. doi: 10.7554/eLife.89828
69
BenoitMBarbaratBBernardAOliveDMegeJL. Coxiella burnetii, the agent of Q fever, stimulates an atypical M2 activation program in human macrophages. Eur J Immunol. (2008) 38:1065–70. doi: 10.1002/eji.200738067
70
YamanakaSSatoYOikawaDGotoEFukaiSTokunagaFet al. Subquinocin, a small molecule inhibitor of CYLD and USP-family deubiquitinating enzymes, promotes NF-kappaB signaling. Biochem Biophys Res Commun. (2020) 524:1–7. doi: 10.1016/j.bbrc.2019.12.049
71
NeklesaTKWinklerJDCrewsCM. Targeted protein degradation by PROTACs. Pharmacol Ther. (2017) 174:138–44. doi: 10.1016/j.pharmthera.2017.02.027
Summary
Keywords
Staphylococcus aureus, macrophage, CYLD, ubiquitin, atopic dermatitis, STAT1, NF-κB
Citation
Schmidt C, Harit K, Traidl S, Naumann M, Werfel T, Roesner LM, Nishanth G and Schlüter D (2025) Ablation of the deubiquitinating enzyme cylindromatosis (CYLD) augments STAT1-mediated M1 macrophage polarization and fosters Staphylococcus aureus control. Front. Immunol. 16:1507989. doi: 10.3389/fimmu.2025.1507989
Received
08 October 2024
Accepted
07 January 2025
Published
28 January 2025
Volume
16 - 2025
Edited by
Patricia Fitzgerald-Bocarsly, Rutgers, The State University of New Jersey, United States
Reviewed by
Tomasz Prajsnar, Jagiellonian University, Poland
Namrata Anand, University of Chicago Medical Center, United States
Updates
Copyright
© 2025 Schmidt, Harit, Traidl, Naumann, Werfel, Roesner, Nishanth and Schlüter.
This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Dirk Schlüter, Schlueter.dirk@mh-hannover.de; Gopala Nishanth, GopalaKrishna.Nishanth@mh-hannover.de
†These authors share senior authorship
Disclaimer
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.